1. Introduction
Chromosome ends, intrinsically prone to DNA damage [
1,
2], are protected by telomeres, repetitive DNA structures stabilized by accessory packing proteins of the shelterin complex [
3]. The telomeric DNA is composed of 5′-TTAGGG-3′ tandem repeat sequences of 5 to 15 kilobase pairs (kbp) [
4,
5] length terminating in a 3′-single-stranded overhang [
6]. Over an individual’s lifetime, somatic cell telomere length diminishes, directly correlating with aging and various diseases through a multitude of distinct mechanisms [
2,
7]. In post-mitotic tissues like the heart, telomere dysfunction, and attrition often coincide with inflammatory processes and other health-threatening disorders [
8,
9]. An established method for assessing changes in telomere length is Quantitative Fluorescence In-Situ Hybridization (Q-FISH). This technique involves fluorescent labeling of telomeric DNA in fixed cells by synthetic peptide nucleic acids (PNA), followed by quantitative fluorescence microscopy [
10].
Q-FISH pioneered the revelation of telomere length heterogeneity within and between clonal cells, offering a valuable tool for quantifying attrition rates [
10,
11]. Initially, it relied on metaphase cell spreads, a labor-intensive method requiring mitotically active cultured cells. However, subsequent studies demonstrated successful applications of Q-FISH in analyzing isolated, post-mitotic cells and tissue sections [
9,
12,
13,
14,
15]. Adaptations for high-throughput screening uncovered substantial variations in telomere lengths among different cell lines and species [
16,
17]. Furthermore, its versatility extends to integration with immunofluorescence and DNA stains, enabling the identification of specific chromosomes, cellular markers, and signaling events [
18,
19].
Despite these achievements, conventional Q-FISH approaches face limitations in fully resolving and analyzing individual telomere structures, necessitating the incorporation of super-resolution techniques.
Super-resolution STimulated Emission Depletion (STED) microscopy [
20] holds tremendous potential, especially in cardiac research, as it is compatible with optically dense, isolated cardiomyocytes or myocardial tissue sections. Despite its recent initiation [
21], this approach has not yet been firmly established in the telomere field and requires specific enhancements for both preclinical and clinical applications. Most Q-FISH studies, including the only available STED study [
22], have intrinsic methodological limitations that can result in limited accuracy of telomere detection and hence quantification. In part, this can be attributed to telomere analysis with single but not multiple focal imaging planes, leading to low telomere detection rates and subsequent statistical inaccuracy. Moreover, existing Q-FISH studies have not explicitly addressed the actual number of PNA probe molecules hybridized to telomere DNA. Telomere length has instead been inferred from comparative calibration measurements [
23] or expressed as localization counts in single-molecule microscopy [
24]. Notably, some Q-FISH studies have demonstrated three-dimensional telomere imaging and thus greatly enhanced detection capability [
25]. However, they have relied on diffraction-limited imaging schemes, significantly limiting their applicability to more challenging samples since closely appositional telomeres would likely remain unresolved, as highlighted in a recent study [
26]. Despite the application of deconvolution procedures in these studies, the axial sectioning capability has remained an additional limit for attaining higher detection efficiencies.
In this study, innovative labeling, recording, and analysis strategies for human telomere samples have been developed. 3D STED nanoscopy has been implemented to separate individual telomeres at high resolution in three dimensions, enhancing geometrical measurements. This includes adaptive illumination, an imaging approach to minimize photobleaching effects [
27]. A corresponding strategy for 3D image analysis is introduced to result in robust and automated measurements at high telomere coverage rates.
Thereby, cell cycle chromatin dynamics were successfully analyzed in a human model system. Based on calibration measurements, the number of hybridized PNA-probe molecules was directly assessed, addressing the relationship between telomere length, compaction, and probe accessibility. Moreover, individual telomeres analyzed by STED could be assigned to specific chromosomes in situ, opening avenues to analyze individual chromosome-specific aberrations. Additionally, initial applications on paraffin-embedded human cardiac samples have been developed, extending this tool to complex cell types such as post-mitotic neurons and muscle cells.
3. Discussion
We established novel FISH workflows with respect to staining diverse samples and introduced new imaging modalities. Specifically, we successfully applied STED microscopy to enhance FISH imaging, focusing on clustered interphase telomeres in HeLa cells with varying chromosome lengths. Expanding our approach, we extended STED-FISH to encompass primary mouse cardiomyocytes and human cardiac biopsies, incorporating dual FISH for in situ chromosome specificity. This extension broadens the possibilities for translational method development and applications, particularly in non-dividing cells such as neurons and muscle cells. This holds special significance, considering the increased attention required for understanding telomere changes in these specific cell types. In the heart, telomere dysfunction causes impaired mitochondrial biogenesis, leading to severe metabolic and contractile dysfunction [
9,
42]. Similarly, telomere attrition has been linked to neuronal dysfunction in neurodegenerative diseases such as Alzheimer’s [
43]. While telomere shortening has been demonstrated to impede the reproductive capacity of stem cells and instigate the onset of age-associated diseases, it’s essential to note that telomere dysfunction and the activation of DNA damage response (DDR) pathways can manifest independently of telomere attrition. This can occur through the loss of shelterin components, whether associated or not with chromatin decompaction [
24,
35,
44]. Importantly, telomere (de-)compaction can be directly quantified using biosensitive structure-function imaging approaches such as STED-FISH.
We enhanced STED-FISH applications through the integration of 3D STED super-resolution microscopy in conjunction with DyMIN adaptive illumination. This innovation yielded unprecedented coverage of individual telomeres in HeLa 1.3 cells and provided highly sensitive readouts.
Regarding PNA probe concentrations, the literature displays variations in the amounts used, influenced by the optimal balance between signal intensities and background. In the Nature Protocol for cardiac Q-FISH [
45], a final concentration of approximately 1.7 µM Cy3-(CCCTAA)
3 PNA (TelC) probe was recommended. For paraffin-embedded cardiac tissues, we utilized 2 µM of the Abberior Star635P or Star Red–labeled PNA probe. In the case of metaphase spreads, HeLa cells, and isolated mouse ventricular cardiomyocytes, we achieved signal intensity saturation at 200 nM of the PNA probe and maintained this concentration. Similarly, Vancevska et al. [
35] employed 100 nM of Alexa647-labeled PNA probe, while a basic Q-FISH protocol [
23] suggests 1 µM of Cy3-(CCCTAA)
3.
The STED-DyMIN modality allowed for minimization and thereby control of the level of photobleaching as a prerequisite to robust, automated image analysis. Additionally, the cell cycle was tracked by concomitant immunofluorescence staining of nuclear speckles. We prefer this approach over the use of synchronized cells since it involves less manipulation of sensitive biological material. The synchronization process can induce stress on cells, and stress responses may introduce variability in experimental outcomes. Moreover, not all cells synchronize uniformly, leading to a mix of cells in different phases of the cell cycle, and cell cycle distribution might require additional control, like flow cytometry. Given our specific research interests, it’s essential to acknowledge that cell synchronization methods were primarily designed for dividing cells, and their applicability to post-mitotic cells, like cardiomyocytes, is limited.
Consequently, the study presented here is the first to investigate telomere changes during the cell cycle using STED nanoscopy combined with Q-FISH analysis. In M-phase, our approach achieved monitoring over 200 telomeres per cell, which was significantly higher compared to all previous Q-FISH studies, including those on HeLa 1.3 cells with elongated telomeres [
16,
28,
41]. The high number results from our increased 3D chromosome coverage and the aberrant genome of HeLa cells, which is hypertriploid with 76–80 chromosomes [
46] in contrast to the 46 chromosomes in healthy human cells. Assuming the same state of polyploidy and chromosome counts for HeLa 1.3 cells, there are putatively 312 telomeres in the M-phase, of which 66% were evidently detected in our data. In interphase cells, assumed to have 156 telomeres in G1, the longest cell cycle phase, a slightly lower detection efficiency of 58% was achieved. Although telomeric clustering could potentially lead to merged spots and thus affect detection efficiencies, it was an unlikely factor in the presented data, as telomere size was precisely resolved and unchanged by the cell cycle stage (
Figure 4D).
Our findings suggest that the analyzed interphase cells were likely in the G1 phase. In contrast, during the G2 phase, doubled chromatids appear, ultimately increasing telomere counts. Importantly, the consistent doubling of spot counts in the M-phase confirms the robustness of telomere detection and argues against spot merging due to spatial crowding or association. Our analysis revealed a homologous population of interphase nuclei with 90 ± 8 telomeres detected by 3D FISH-STED (mean ± SD). Hence, our analysis did not indicate widely different ploidy in HeLa 1.3 cells, with a consistent doubling of spot counts in M-phase (n = 207 ± 19), showing low relative variance between cells in both groups (CV < 10%). Therefore, we can rule out variable polyploidy as a source of error in this study. Another potential source of error could be a large fraction of interphase nuclei in G2, where telomere length measurements and telomere counts might be affected by the ongoing synthesis of new telomeric DNA. As noted above, we detected smaller variability in telomere counts in the S versus M phases, arguing against a relevant fraction of nuclei in G2. In parallel, we detected nearly equal telomere sizes (
Figure 4D) in the S and M phases, arguing against the presence of potentially decompacted, enlarged G2 telomeres during synthesis. This is further supported by the poor correlation of telomere volumes to signal density (
Figure S2), while a high correlation would reflect better probe binding during decompaction below. In addition, Deplanche et al. [
47] showed that asynchronous HeLa cells are 79% in G1-, 17% in S, and only 14% in G2/M-phase. This might equally hold true for HeLa 1.3, though it has not been studied to our knowledge. Whether telomere elongation is a sufficient prerequisite for elongated G2 phases is likewise unknown. Therefore, we believe we would have distinguished G1 versus G2 phase nuclei.
In our study, a cell-cycle-dependent variation of telomere size could not be reproduced, contradicting previous PNA-FISH-STORM data and the telomere decompaction model [
24]. Our method reported an average telomere size of 132 nm (FWHM), corresponding to a mean volume of 1.44 × 10
−3 μm
3 when assuming a nearly spherical shape of telomeres, consistent with previous studies [
24,
35]. These values are comparable to previously reported data, despite different super-resolution techniques, e.g., PNA-FISH-STORM in HeLa cells, measuring telomere size in terms of convex hull volume (5 × 10
−3 μm
3, HeLa 1.3) and gyration radius (88 nm in HeLa L, a HeLa subclone with elongated telomeres).
Bandaria et al. [
24] revealed a dramatic increase in telomeric volume after the knockdown of specific shelterin proteins, termed decompaction, which coincided with local DNA-damage response (DDR) signaling. This confirmed the protective and structural functions of the shelterin complex and proposed a functional connection. Moreover, their study found the telomeric volume to be dependent on the cell cycle and proportional to telomere length, demonstrating telomere compaction in metaphase. However, later STORM studies in murine [
44] and HeLa cells [
35] contradicted the decompaction model, as a telomere volume increase was not detected after Shelterin knockdown, despite the appearance of DDR signals on dysfunctional telomeres. Thus, the DDR-protective function of Shelterin proteins has been confirmed and further elaborated by FISH nanoscopy, while the role and dynamics of telomeric chromatin (de-)compaction remain controversial.
Surprisingly, a decrease in the peak and total brightness of telomere spots by 21% was observed in the M-phase without significant changes in telomere size, leading to a decrease in signal density at the single telomere level. This finding contradicts the hypothesis of an increased signal density upon chromatin compaction in metaphase. Telomere brightness, measured by the integrated signal of telomere spots, directly represents the number of FISH probes bound to telomere sequence repeats. The overall telomere sequence length was considered constant within the same population of cultured cells and the lateral size of telomere spots remained constant. Therefore, it seems plausible that the molecular accessibility of telomeric DNA for FISH probes was reduced in the M-phase of the mitotic process, thus lowering the labeling efficiency at a molecular level. This finding confirms increased methodological variability of in situ Q-FISH compared to metaphase Q-FISH [
16], which can be explained by dynamic protein recruitment [
48] and chromatin remodeling [
49,
50] happening at telomeres under the control of the cell cycle [
51,
52].
In the recent FRET-FISH study, however, the integration of oligo DNA-FISH and FRET allowed for the measurement of chromatin compaction at individual gene loci in single cells across different cell cycle phases. The study specifically targets the biophysical properties associated with the interplay of chromatin accessibility and compaction state. Importantly, the efficiency of oligo hybridization in FRET-FISH was not compromised, even when targeting the highly condensed chromatin during mitosis [
53].
In contrast, our findings challenge a foundational assumption of the PNA-Q-FISH method, which relies on a linear conversion of fluorescent brightness to telomere length, assuming a constant and high labeling efficiency of PNA nucleotides. The data presented here provide compelling evidence for a low labeling efficiency ranging from 10% to 20% (
Figure S3) that varied with the cell cycle. Absolute telomere lengths derived by commonplace calibrations of Q-FISH should therefore be interpreted with care, as calibration systems are inherently different from the biological samples at hand.
In conclusion, the main findings and novelties of the present work are, firstly, the evidence of low labeling efficiency and variable coverage of telomeric tandem repeats by PNA-FISH probes compared to telomere lengths confirmed by Southern blot TRF analysis. Secondly, we overcame the intrinsic technical limitations of conventional 2D approaches by utilizing 3D-STED-DyMIN Q-FISH, where individual telomeres were detected and quantified at the appropriate imaging plane, overcoming focal shift problems. The high coverage of telomeres obtained confirms the strength of our 3D imaging and analysis approach, resulting in a more reliable and accurate readout of individual telomere brightness. Additionally, STED imaging confirmed sufficient spatial telomere separation in the xy-plane. Hence, we present a novel high-resolution tool for nanometric imaging of cardiac cells and tissue samples. This allows for the future study of telomere integrity in situ with key model systems and human samples from cardiovascular and neurodegenerative diseases, potentially in the context of future therapeutic strategies.
4. Materials and Methods
4.1. Cell Culture
Human cells were cultured at 37 °C in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FCS, Gibco) and antibiotics.
4.2. Human Specimens
Following informed consent, small left-ventricular endomyocardial biopsies were obtained from two patients undergoing aortic valve replacement at the University Medical Center Göttingen [
54]. Left-ventricular biopsies were directly fixed in 4% paraformaldehyde, embedded in paraffin, and cut into 5 μm sections for the imaging study.
4.3. Isolation of Cardiac Myocytes from Mouse Ventricles
Ventricular myocytes were isolated from 8–20-week-old wild-type C57BL6/N mouse hearts, as previously described [
55]. Briefly, mouse hearts were excised, mounted via the aorta to a cannula connected to a modified Langendorff perfusion setup, and perfused with a modified Ca
2+-free Tyrode buffer (120.4 mM NaCl, 14.7 mM KCl, 0.6 mM Na
2PO
4, 0.6 mM KH
2PO
4, 1.2 mM MgSO
4, 10 mM HEPES, 4.6 mM NaHCO
3, 30 mM Taurin, 10 mM 2,3-Butanedione monoxime, 5.5 mM Glucose, pH7.4) at 37 °C. The hearts were then enzymatically digested by adding 2 mg/mL collagenase type II and 40 µM CaCl
2 to the Tyrode perfusion buffer for 9 min at 37 °C. The ventricles were dissected and agitated in solution to release the cardiomyocytes. Finally, the isolated ventricular cardiomyocytes were transferred into Tyrode buffer containing 10% FBS (Sigma-Aldrich, Burlington, MA, USA) to terminate the enzymatic digestion.
4.4. FISH Procedure
Sample Preparation
Metaphase spreads were prepared according to a published protocol [
23]. In brief, Hela cells were subcultured 36–48 h before colcemid treatment, aiming for approximately 70% confluency at the time of treatment. Subsequently, the cells were incubated for 15 h in a regular medium containing 0.1 μg/mL of colcemid to arrest them at metaphase and harvested by trypsinization.
After hypotonic swelling (15 min, 37 °C) in prewarmed KCl solution (0.025 M sodium citrate and 0.04 M KCl), cells were fixed in cold Carnoy fixative (3:1 (v/v) methanol/acetic acid) for 10 min at RT. The fixation process was repeated twice, then cells were washed in a fixative solution, dropped onto ice-cold glass slides, and dried on top of a humidified heating block (1 min, 80 °C).
Alternatively, HeLa cells, cultured on collagen-coated microscope slides up to 60–70% confluency, were fixed in 4% formaldehyde in PBS for 10 min at RT. The cells were then washed twice with PBS and permeabilized in Triton X-100 buffer (0.5% Triton X-100, 20 mM Hepes-KOH (pH 7.9), 50 mM NaCl, 3 mM MgCl2, 300 mM Sucrose) for 10 min at RT. Subsequently, they were fixed again in 4% formaldehyde in PBS for 10 min at RT.
Mouse ventricular cardiomyocytes (freshly isolated as described below) were plated on glass slides coated with mouse laminin (BD Biosciences) and allowed to settle for 20 min at RT. Adherent cells were consequently fixed with 4% formaldehyde in PBS for 10 min at RT and processed as described above for HeLa cells.
Alternatively, nuclei were isolated from mouse ventricular cardiomyocytes using the Nuclei Isolation Kit (Nuclei EZ Prep, Sigma-Aldrich), fixed with formaldehyde, and processed as described above.
The prepared slides were utilized for FISH studies or stored at −20 °C for future use.
The slides with the attached samples were rehydrated in PBS and fixed with 4% formaldehyde in PBS. After washing, the slides were treated with 100 μL of RNAse A solution (100 µg/mL of RNAse A in 2× Saline Sodium Citrate (SSC) buffer) for 1 h at 37 °C in a moisture-sealed slide incubation chamber. Then they were washed three times in 2× SSC and once in distilled water. Subsequently, slides were immersed in 0.005% pepsin solution (0.005% pepsin in 10 mM glycine, pH 2) for 4 min at 37 °C. After washing in PBS, the slides were fixed once more in 4% formaldehyde, washed in PBS, and dehydrated at RT in an ethanol series (70%, 85%, and 100%).
Subsequently, the slides were hybridized with Abberior Star 635P- or Abberior Star Red-labeled PNA probes (TelC, 2 µM or 200 nM) in a hybridization mix (22 mM Na2HPO4, 22 mM Tris-HCl, pH 7.4, 66% formamide, 2.2×SSC, 0.11 µg/mL Salmon Sperm DNA). TelC with N-terminal Star-635P or Star Red bound via two AEEA linker elements was custom synthesized by Eurogentec (Eurogentec Ltd., Koeln, Germany). DNA was denatured for 10 min at 85 °C, and hybridization was carried out in a moisturized hybridization chamber at 4 °C overnight or at RT for three hours.
For duplex FISH, 40 ng of Green 13q-ter subtelomeric probe (Oxford Gene Technology, CytoCell Ltd., Cambridge, UK) was added to the hybridization mix together with the TelC probe or alone as a control, and specimens were denatured and hybridized as described above.
After hybridization, the slides were incubated twice in a washing solution (2×SSC, 0.1% Tween-20) for 15 min at 60 °C. They were then washed twice in 2× SSC at RT and once in distilled water. Slides were air-dried, protected from light, and mounted with Prolong Gold mounting media with DAPI (Invitrogen).
4.5. IF-FISH Procedure
HeLa 1.3 cells were plated on collagen-coated coverslips and allowed to attach for 24–36 h before fixation, resulting in 60% confluency. After washing with PBS, cells were fixed in 4% formaldehyde in PBS for 10 min at RT. Following fixation, cells were washed with PBS and permeabilized in Triton X buffer for 10 min at RT. After permeabilization, cells were washed with PBS and blocked with a blocking solution (1 mg/mL BSA, 3% goat serum, 0.1% Triton X-100, 1 mM EDTA; PBS pH 8.0) containing 100 μg/mL DNAse-free RNAse A for 30 min at 37 °C. After blocking, cells were incubated with primary antibodies (anti-SC-35, 1:200, mouse monoclonal, Abcam) for 2 h at RT in blocking solution and washed in PBS three times for 5 min. Afterwards, cells were incubated with secondary antibodies (Abberior Star-580, 1:200) in a blocking solution for 2 h at RT and washed in PBS three times for 5 min.
For the following FISH procedure, cells were fixed once again in 4% formaldehyde in PBS for 10 min at RT, washed in PBS, and sequentially dehydrated in 70%, 85%, and 100% ethanol, respectively. DNA denaturation and subsequent hybridization were performed as described above.
4.6. FISH Procedure for Paraffin-Embedded Cardiac Tissues
The protocol was adapted from Sharifi-Sanjani et al. [
45]. Briefly, the paraffin on the slides was melted by incubating them in a slide warmer at 65 °C for 5 min. For deparaffinization, the slides were incubated twice in fresh xylene for 7 min each at RT. Slides were then rehydrated in an ethanol series (100%, 95%, and 70%), then in distilled water at RT, followed by rinsing in 1% Tween-20 and distilled water. Afterwards, the antigen retrieval procedure was performed. Antigen retrieval was conducted by incubating the slides in a 1/10 dilution of antigen retrieval citric buffer (Sigma-Aldrich) at 95 °C for 45 min in a water bath. Subsequently, the slides were dehydrated in an ethanol series (70%, 95%, and 100%) and air-dried.
DNA denaturation and hybridization steps were carried out as described above, with the final concentration of the PNA probe set at 2 µM. The denaturation step for human samples was performed for 5 min at 85 °C. Following hybridization, the slides underwent two 15 min washes in 70% formaldehyde, 10 mM Tris-HCl, pH 7.5 at RT in the dark. After drying in the dark, the slides were mounted as described above and imaged or stored at 4 °C in the dark.
4.7. TRF Assay
DNA was extracted from cells using the DNeasy Blood and Tissue Kit (Qiagen) following the manufacturer’s instructions. DNA concentration and intactness were assessed using the Genomic DNA Screen Tape (Agilent Technologies) on the 2200 Tape Station (Agilent Technologies). The southern blot was performed using the TeloTAGGG Telomere Length Assay (Roche), following supplier instructions with a few changes. Then, 500 ng of DNA was digested for 5 h. The southern blot was analyzed using the software Image Quant TL (GE Healthcare, Chicago, IL, USA) and Microsoft Excel, as suggested by Lincz, et al. [
56]. The mean telomere length was calculated with the formula
.
4.8. Optical Device Setup
All imaging was performed using a custom-built microscopy setup based on the Abberior RESOLFT QUAD P microscope kit. The setup uses an Olympus IX83 inverted microscope, equipped with a 100x 1.4 NA oil-immersion objective and an Abberior QUAD beam scanner. The setup was controlled by the software Abberior Imspector version 16.1.6905. Three fluorescent imaging channels were used in confocal scanning mode, herein designated as blue (405 nm excitation, 422–467 nm detection), green (594 nm excitation, 605–625 nm detection), and red (640 nm excitation, 650–720 nm detection). All lasers were pulsed and regulated by an acousto-optic modulator. The red channel was also used in STED mode, supported by a 775 nm synchronized depletion laser, which was aligned at the start of each imaging session using 40 nm red fluorescent beads (Abberior Nanoparticle Set for Expert Line 595 and 775 nm). In STED mode, a time gating of 0.5–8 μs was used. The pinhole size was set to one Airy unit in all recordings. Reported pixel intensities are photon counts detected by avalanche photodiode detectors without further processing. The intensity calibration workflow that was demonstrated is nonetheless applicable to a variety of setups that report arbitrary intensity units, as long as the photon-to-brightness response is linear, i.e., detectors are not reaching saturated conditions. This can be evaluated based on the linear fit, as shown in
Figure 6C.
Z-stacks were recorded using continuous autofocus, an operation mode used to mitigate sample drift by compensation of small stage movements along the optical axis.
4.8.1. Dynamic Intensity Minimum STED Microscopy
IF-FISH slides of HeLa 1.3 cells were prepared according to the customized protocol shown below and imaged in three channels: DAPI staining in blue, nuclear speckles IF in green (immunostaining using sc35 primary antibody, Abberior Star-580 secondary antibody), and telomere FISH in the red channel. Confocal overview images containing multiple cells were recorded at 100 nm pixel size in the blue, green, and red channels. The respective total pixel dwell times were 20, 60, and 20 μs and laser powers were 25, 20, and 2%. From the overview images, the M-phase and interphase cells were identified and chosen in random order for 3D-STED-based acquisition.
The 3D-STED imaging of single nuclei for advanced Q-FISH telomere analysis was performed with a setting of 25% 3D-STED. Images were recorded using 30 × 30 × 200 nm pixel size (x, y, z), 8% excitation intensity, and 20% STED intensity using Dynamic Intensity Minimum (DyMIN) STED mode [
27]. DyMIN parameters were set as follows: CONF level and DyMIN level were set to 15 counts. The base pixel dwell time was 10 μs (5 μs × 2 line accumulations), with one step in the confocal channel, 2 steps in the probing channel, and 4 steps in the final STED channel, resulting in a dwell time of 40 μs in the recorded STED image. The Exclusion parameter was set to 10, leading to a probing step STED intensity of 6.3%, or about one-third of the final power.
4.8.2. Molecular Brightness Calibration
To retrieve fluorophore counts from fluorescent photon counts, fluorescent bead slides (Custom Brightness DNA Origami, GATTAquant) with defined fluorophore counts of 17, 34, or 60 molecules of Star-635P per bead were measured to create a calibration curve [
57]. Bead samples were produced using the same mounting medium as biological samples (ProLong Gold Antifade, Thermo Fisher Scientific, Waltham, MA, USA).
4.8.3. Image Analysis
In ImageJ Fiji version 1.52s38, microscopy images were manually cropped to include single nuclei and masked by polygons following the DAPI signal to exclude background signals if necessary. Display range and color maps were adjusted to validate the fitting process and present the data. The raw intensity values were not changed by these operations, and files were saved in the uncompressed 16-bit “.tif”-format.
In Q-FISH analysis, TelC-Star635P signals were smoothed by a Gaussian filter and then subjected to a 3D spot detection routine based on the ImageJ plugin software FindFoci, implemented within a custom macro workflow [
33]. A background parameter of 10 photon counts and a minimum spot size of 10 voxels were determined empirically using the provided graphical user interface. The resulting center of mass coordinates for each telomere spot were used to center two orthogonal lateral fits of the signal by a Gaussian function using the ImageJ curve fitting tool. The Gaussian function in ImageJ is defined as follows:
in which
a is the y-axis-offset or background level,
b is the signal amplitude,
c is the x-axis-offset and
d is the standard deviation (σ). A line length of 480 nm (16 px) was used in STED images. The fit with higher goodness (r²) was used to retrieve telomere brightness and lateral size by the following equations when at least one fit was sufficient (r² > 0.8):
Volume was approximated by a 3D Ellipsoid, in which radius was defined as half of FWHM and was interpolated in z by the average of x and y because lateral resolution was much higher than axial resolution. If one fit was insufficient, only the other fit was used.
Further spot statistics were only determined when both fits were sufficient. Integrated brightness was calculated using the 2D Gaussian integral of a spot. Signal density was calculated as the ratio of integrated brightness and volume.
4.9. Data Handling and Statistics
Data visualization and statistical analysis were performed in GraphPad Prism version 8.3.0. Scatter plots show raw data retrieved by automated spot analysis, with red lines indicating the median and interquartile range. Larger datasets, such as combined data from multiple cells, were displayed as box plots showing the median and interquartile range, with whiskers denoting the 5th and 95th percentiles. Bar graphs displayed the average n (telomere number per cell) ± SD.
Statistical significance was determined using a nested/hierarchical one-way ANOVA or
t-test. In this test design, cell origin or cell cycle state were used as higher-order groups and single cells as subgroups. Combining the data of multiple cells in one group and then using standard tests is common practice even in Q-FISH studies, but results in pseudo-replicates and thus falsely low
p-values [
58], since there may be high variability between cells, meaning that data points are not truly independent. The nested tests, however, can account for unequal sample sizes and variance both across subgroup data and subgroup means, which are accounted for in the group comparison. Since the test assumes a Gaussian population, data following lognormal distributions, such as brightness values, were transformed in Prism by the formula Y = log
10(X) prior to statistical testing.
Here, p values below 0.05 were defined as significant and annotated in graphs as follows:
ns = p > 0.05, * = p ≤ 0.05, ** = p ≤ 0.01, *** = p ≤ 0.001
Frequency distribution data were fit by Gaussian curves of the following formula:
using robust regression and the retrieved μ and σ values used as the sample mean and SD. Lognormal distributions were similarly modeled based on the following equation:
using least-squares regression. Multiple fits were compared using the extra-sum-of-squares F test, testing if one model can fit all datasets.